Introduction

The Polymerase Chain Reaction (PCR) is used in all areas of biological science research, including the clinical, forensic and diagnostic fields and the widespread adoption of the PCR technique has revolutionized life science research.

Many improvements have been made to the basic technique and many adaptations have evolved, including reverse transcription PCR (RT-PCR), quantitative real-time PCR (qPCR), reverse transcription qPCR (RT-qPCR) and digital PCR (dPCR). This guide is designed for scientists with a background in molecular biology and for scientists in other fields who are interested in learning more about PCR-based techniques. Basic principles are presented and supplemented with practical examples and theoretical concepts.

To illustrate the theoretical concepts, protocols are provided that can be used to analyze sample quality, optimize assay conditions, determine assay efficiency and RT linearity. These protocols can be easily adapted for all research applications.

Historical Timelines

In 1971, Kjell Kleppe and the Nobel laureate, Gobind Khorana, published a description of a technique that represented the basic principles of a method for nucleic acid replication:

“… The DNA duplex would be denatured to form single strands. This denaturation step would be carried out in the presence of a sufficiently large excess of the two appropriate primers. Upon cooling, one would hope to obtain two structures, each containing the full length of the template strand appropriately complexed with the primer. DNA polymerase will be added to complete the process of repair replication. Two molecules of the original duplex should result. The whole cycle could be repeated, there being added every time a fresh dose of the enzyme. It is however, possible that upon cooling after denaturation of the DNA duplex, renaturation to form the original duplex would predominate over the template-primer complex formation. If this tendency could not be circumvented by adjusting the concentrations of the primers, clearly one would have to resort to the separation of the strands and then carry out repair replication. After every cycle of repair replication, the process of strand separation would have to be repeated.”1

While this passage appears to be a clear description of the process that is now recognized as the PCR, it could not be verified experimentally at the time because the target sequences required for primer design were not readily available, nor was there a convenient method for manufacturing the synthetic PCR primer oligonucleotides.

Originally, when considering the process of DNA amplification, Kary Mullis, et al.,2 had assumed that when primers were added to denatured DNA, they would be extended, the extension products would become unwound from their templates, be primed again and then the process of extension repeated. However, this was not the case and the DNA had to be heated almost to boiling after each round of synthesis to denature the newly formed, double-stranded DNA. The Klenow fragment of DNA polymerase I that was being used for synthesis was then inactivated by the high temperature and so more enzyme was required at the start of each cycle, as predicted by Kleppe1. A critical development that led to the universal adoption of the PCR technique was the concept of using a thermal stable DNA polymerase that could tolerate the high temperature of the repeated denaturation steps. The Klenow fragment of DNA polymerase I was replaced with the heat tolerant Taq DNA polymerase3,4. When using Taq DNA polymerase, several rounds of amplification could be executed in a closed reaction tube without replenishing the enzyme. In addition, larger fragments could be amplified, the use of higher temperature reactions was sufficient to increase replication fidelity, reduce nonspecific product formation and allow the products to be detected directly on ethidium bromide stained, agarose gels5,6,7.

The 1993 Nobel Prize for Chemistry was awarded jointly to Michael Smith “for his fundamental contributions to the establishment of oligonucleotide-based, site-directed mutagenesis and its development for protein studies” and to Kary Mullis “for his invention of the polymerase chain reaction (PCR) method.”

At around the same time that the 1993 Nobel Prize for PCR was awarded, Higuchi, et al.,8 recognized the process of PCR could be monitored by addition of a fluorescent label that binds to the accumulating PCR product. As the concentration of PCR product increases, the intensity of the fluorescence signal also increases. This discovery paved the way for modern quantitative real-time PCR (qPCR). In current qPCR technology, these fluorescence signals are generated by inclusion of either fluorescent DNA-binding dyes or oligonucleotide probes. Fluorescent DNA-binding dyes, such as SYBR® Green I, are included in the PCR buffer. When the dye is free in solution, it emits excess energy as vibrational energy. However, as the DNA target is amplified, the dye binds to the DNA product and adopts an alternative conformation. This conformational change reduces the molecular mobility and results in the excess energy being emitted as fluorescence. Hence, the DNA-bound dye has a higher fluorescence than the unbound dye. An alternative approach to monitoring the reaction is to include a labeled primer in the reaction9 or, for additional specificity, an additional oligo probe is situated between the two primers. This oligo probe is labeled and in most cases, also quenched. Various probe options are available, but the most popular are the Dual-Labeled Probe (also referred to as TaqMan®)10, Molecular Beacons11, Scorpions® Probes12 and LightCycler® Probes13 (see Quantitative PCR and Digital PCR Detection Methods).

The most recent qPCR development has been digital PCR (dPCR) in which a sample is diluted and divided into hundreds to millions of reaction chambers. The dilution and partition must be such that each chamber contains either zero or one template copy and the starting concentration is estimated from the ratio of partitions with a positive signal to those with a negative signal. A Poisson distribution is used to quantify the target copies. Since the target is diluted to a single copy, dPCR is particularly useful for applications requiring quantification of rare mutations amidst a high concentration of wild-type sequences (see Digital PCR).

In addition to genomic DNA sequence analysis, it has long been believed that investigating specific mRNA sequences with PCR can yield essential information about the biology of the cell. Studying gene quantity changes between normal and diseased tissues, or looking for changes in gene expression in response to drug treatments, is being used to understand how regulation of gene expression is a part of the complex system of cellular control. Measuring mRNA requires an additional reverse transcription (RT) step in order to convert RNA to a DNA template that is suitable for PCR amplification. This is carried out using a reverse transcriptase enzyme and extension from one or more oligo primers. Priming of the reverse transcription reaction may be from a sequence specific primer, from a series of random primers that hybridize along the length of the mRNA, or from a primer directed towards a tract of adenosines that are present on the 3’ end of most messenger RNA sequences (polyA tail). After elongation from the primer, a double-stranded hybrid of RNA and DNA, called first-strand cDNA, is produced. This cDNA is then a suitable template for PCR and relative quantities of specific RNA templates are determined by semi-quantitative or preferably quantitative PCR (see Reverse Transcription).

More recently, a family of small, non-coding RNA (ncRNA) species, including microRNA (miRNA), has been identified. It is increasingly apparent that the expression profile of miRNA molecules can be used as genetic biomarkers that are characteristic for specific diseases. Using PCR to amplify these targets is particularly challenging because they are very small and do not have natural polyA tails. However, there are options for analyzing the genetic information of these molecules, such as capturing all the miRNA in a sample by adding an artificial polyA tail and then performing selective qPCR of the target of interest (see PCR/qPCR/dPCR Assay Design).

Regardless of the chosen PCR technique or application, in theory, a single template molecule should be replicated during each PCR cycle. Assuming absolute, perfect replication at each cycle, this would lead to 240 or 1012 identical molecules (amplicons) after 40 rounds of amplification. However, the actual sensitivity of any PCR-based assay is determined by the assay design14, as described in PCR/qPCR/dPCR Assay Design, sample quality and the susceptibility of the assay to inhibitors, as described in Sample Purification and Quality Assessment, and optimization, described in Assay Optimization and Validation. A functioning assay requires that all of these factors be considered prior to application.

Along with dynamic range, the analytical sensitivity of a PCR-based assay depends on the variability of the data. The presence of inhibitors in the sample matrix, degradation and unspecific amplification of off-target products in the reaction are all potential sources of variability. If these occur randomly, or pseudo-randomly, in the biological samples, measuring replicates can minimize the impact of the variability on the end result. However, there may be cases where the variability is not random, but systematic. In these cases, averaging of replicates will not alleviate the problem. For both systematic and random technical errors, experimental approaches to minimize the errors are preferred.

In addition, to protect against uncertainties due to reaction variability, it is important to include a series of controls alongside all samples in any experiment. The choice of controls and whether to include them should always depend only upon the nature of the study. Scientific integrity should be the driving factor behind all experimental design. It amounts to a false economy of all resources to run inadequate experiments that could even result in retractions15,16. Some controls should certainly be considered obligatory and original data should be presented for inspection, especially when the results of the study or analysis may lead to life and death decisions17. Controls that are run in parallel with experimental samples are widely used as verification of assay quality and during the troubleshooting process, as described in Troubleshooting. In recent years, the importance of reporting assay quality has been recognized. A team of internationally renowned, expert Molecular Biologists compiled The MIQE Guidelines: Minimum Information for Publication of Quantitative Real-Time PCR Experiments17. These guidelines direct the researcher through the process of assay quality control. Information derived from controls during assay verification is required to establish that the experiment is MIQE compliant. As a follow up to the publication of these recommendations, a review of qPCR publications was made and an increase in adoption of the guidelines was shown18. Similar recommendations for studies containing data derived from dPCR experiments were published recently19.